We developed a sensor to target antibiotic resistances in E.coli to test if a specific strain carries a certain antibiotic resistance gene. Our sensor system consists out of a phagemid with a CRISPR/Cas system and LacZ as a reporter under the control of a pRECA promoter (SOS response promoter).
If our CRISPR/Cas system can bind to the target (antibiotic resistance gene), the Cas9 generates at this specific target site a double strand break, which starts the expression of our reporter, as the promoter gets active at stress resulting from double strand breaks (Figure. 1). Having the system on a phagemid, the sensor system will spread all over a population, to get a clear color output if the target has been detected. Depending on target sequence the system carries, we can identify different antibiotic resistances in a strain. This is a novel approach of detecting genes in bacterial strains. We used E.coli and a M13 phagemid to target the kanamycin resistance gene.
Figure 1. The CRISPR/Cas system generates a site-specific double strand break in the kanamycin resistance gene. This break activates the expression of the reporter and a color output can be seen.
This sensor is a proof of concept for a similar system in mycobacterium tuberculosis. Such a system could potentially be used to test if a patient has TB and what type of resistance genes the specific strain contains to adapt the patient’s drug treatment.
Motivation and existing TB detection systems
Tuberculosis (TB) remains a major global health problem. While the treatment of this disease in countries with adequate medical care is fairly easy to detect and treat, it remains hard to treat and diagnose in poorer countries.
Up to now, a high quality lab that uses modern diagnostics is a prerequisite for early, rapid and accurate detection of TB. Therefore, diagnosis of TB and drug resistant TB remains a particular challenge for laboratory systems, especially in developing countries. The lack of cheap, quick and accurate tests make it hard to control the Tuberculosis epidemic, which claims millions of lives every year in developing countries. Setting up a cheap, fast and culture-based method could therefore decrease diagnostic time and facilitate patient treatment.
The most common method for diagnosing TB nowadays is sputum smear microscopy, in which bacteria are observed in sputum samples of patients under the microscope. However, this cannot be used to identify paucibacillary (containing just a few bacteria) or extrapulmonary (outside of the lungs) TB. Diagnosis methods using culture methods require laboratory infrastructure that is not widely available in countries with a high burden of TB and results are only available after a few weeks.
Other conventional methods used to diagnose multidrug-resistant TB (MDR-TB) also rely on the culturing of specimens followed by drug susceptibility testing (DST). Results take weeks to obtain and not all laboratories have the capacity to perform DST of first-line or of second-line drugs. MTB/RIF is a new rapid molecular test that can diagnose TB and rifampicin-resistant TB within hours. Molecular tests, such as GeneXpert, unlike culture-based methods, are fast, accurate and can detect drug-resistant strains. But the high costs and need for laboratories make access an issue for developing countries. The new method, published in the Journal of Applied Microbiology, uses a microcalorimeter to detect heat produced by Mycobacterium tuberculosis, the bacterium that causes TB, on a growth medium. The study showed that detection takes 4–5 days but more sensitive microcalorimeters could detect tuberculosis in 24 hours.
System Design
To use our system, we need two strains, a phagemid producing strain and the target strain. The producer strain contains a helper plasmid, which produces the capsid proteins for the phage and the phagemid plasmid with the sensor elements (Figure 2), which is packed up into the M13 capsids (Figure 3). The helper plasmid encodes for everything of the phage but doesn’t contain the packaging sequence, while the phagemid plasmid only contains the packaging sequence but doesn’t code for anything else of the M13 phage. As M13 is a lysogenic phage, the phagemid particles are exported into the media, where they can be collected (Figure 4) and transferred to the target strain (Figure 5). The phagemid infects cells that are conjugating, as M13 needs a sex pili to attach to the cell and release the plasmid plasmid into the cell (Figure 6). Within the target strain, the Cas9 protein as well as the trRNA and crRNA are expressed (Figure 7). The crRNA and trRNA are further processed and form a hybrid that attaches to the Cas9 protein (Figure 8). The RNA hybrid guides the Cas9 to the target sequence (Kanamycin resistance gene) as it can be seen in Figure 9 and the Cas9 protein generates a target specific double strand break (Figure 10).
This double strand break activates the SOS response promoter pRECA (Figure), which then starts the expression of β-galactosidase (Figure 11). In the presence of xgal, xgal is hydrolyzed by β-galactosidase, which gives a blue color output (Figure 11 and 12).
Background
CRISPRs
The mechanism we are using for our sensor is based on the CRISPR/Cas System. The CRISPR/Cas genome editing method, which recently became very popular, is derived from the bacterial “immune system”. CRISPRs are Clustered Regularly Interspaced Short Palindromic Repeats that are part of the bacterial genome. These loci contain multiple short repeats. In between the repeats are so called spacers that are sequences derived from extrachromosomal DNA, e.g. from invading viruses or plasmids. Within bacteria, those CRISPRs are naturally used to detect and destroy foreign DNA that is saved in the spacers.
In total the CRISPRs and the spacers form a so-called CRISPR array that is transcribed as one unit. CRISPR associated proteins (Cas proteins) are involved in further processing and action steps of the CRISPR system. There are many different regulation systems, here we describe the Cas9 system that will be used for our purposes.
The transcribed mRNA is processed into so called crRNA (CRISPR RNA), which includes the spacer sequence of foreign DNA. Together with a transactingCRISPR RNA (tracrRNA), the crRNA forms a duplex that is cleaved by RNaseII. The resulting hybrid serves as guide for the Cas9 protein that generates double strand breaks at the position the RNAs guide it to. By this double strand break the invading DNA is destroyed.
During the last years, researchers discovered the CRISPRs as a method for genome editing. By designing the spacer sequence, specific sequences in the genome can be targeted. With the provision of a sequence with homologous sequences, easy insertion can be done. This method can be used to insert mutations or new sequences nearly everywhere in the genome.
Recently many papers have been published for genome editing in bacteria (Jinek et al., 2012, Jiang et al., 2013), yeast (DiCarlo et al., 2013) and mammalian cells (Mali et al., 2013, Cong et al., 2013). They describe how to use the CRIPRs for different purposes. One requirement to target a sequence is a NGG at the end of the sequence. This NGG sequence is called PAM - protospacer adjacent motif, while the target sequence itself is called the protospacer, which should have a length of around 12 to 20 bp.
In our approach we test two different systems, the systems of DiCarlo et al. 2013 and Jiang et al. 2013. The system of Jiang is a system developed for bacteria and especially for E.coli. The system of DiCarlo is based on a paper of () and adapted for yeast.
Different from the Jiang paper, the DiCarlo paper uses a gRNA (guideRNA) to guide the Cas9. The gRNA is actually the RNA that results after transcribing and folding is the same complex as you get after the processing of the tracrRNA with the crRNA. It is hence an improvement, which makes the design and expression easier. But as the system hasn’t been tested before in bacteria, we will use both systems to make sure we get the desired results.
RecA promoter
Our reporter system consists out of the RecA promoter and LacZ. Originally, the RecA promoter of Escherichia coli has its main role in the activation of the SOS repair system. It is regulated by the LexA repressor, which binds to the SOS box sequence of the promoter. RecA expression is regulated in a way that the number RecA proteins is maintained between 1 000 and 10 000 monomers per cell. DNA damage leads to an inducing signal, which then activates the RecA protein. In the SOS response the LexA repressor is cleaved by the RecA protein, so the full RecA expression can be reached. The RecA protein can then repair both single stranded and double stranded DNA breaks.
In our project RecA promoteris used to detect double stranded DNA breaks in the region of antibiotic resistance genes caused by the CIRSPR/Cas system. To get the desired activation of the promoter, it is important to assure that no other stressing agents might activate the RecA promoter. Those agents could be UV light, X-ray, ionizing radiation and different types of DNA breaking compounds. The SOS response in E. coli could also get induced by engineered M13 phage infection that are defective in the minus-strand origin, and so unable to form the double stranded replicative stage.
In this case the single-stranded DNA would be the SOS-inducing signal. The wild type M13 phage doesn’t induce the SOS response (Higashitani et al., 1992). In another study genomes of phage M13 infected and uninfected E. coli strains were compared by oligonucleotide microarrays, where no stress response genes were scored as upregulated (Karlsson et al., 2005). Regarding the phagemids, an A UV-damaged oriC phagemid did not induce SOS response in a recipient cells oriF phagemids on the other hand did (Sommer et al. 1991). To make sure that our system functions reliably, we will test the activation of the RecA promoter at different stress inputs (UV, phage infection,…) by measuring the fluorescence of YFP driven by the RecA promoter.
LacZ
As a reporter we are using LacZ in our system. LacZ is the part of the lac operon in E.coli and its gene product in an enzyme, the β-galactosidase. This enzyme has 3 catalytic functions: to cleave lactose into glucose and galactose, catalyze the trangalacosylation of lactose to allactose and the cleavage of allactose into monosaccharides (Juers et al., 2012). X-gal (5-bromo-4-chloro-3-indoyl-b-D-galactopyranoside) contains a galactose fused to a substituted indole and is a very common used compound in Molecular Biology. It is colorless, soluble and non-toxic to cells. The β-galactosidase recognizes the galactose of the molecule and hydrolyzes the X-gal. In this process the indole is released which dimerizes. The dimerized indole is insoluble and has a blue color. Due to this color output it is often used to confirm cloning. Only small numbers of enzyme and X-gal are needed to actually get a color detection, which makes it very attractive for our system.
Literature (not complete yet)
Higashitani N., Higashitani I.A., Roth A., Horiuchi A.K. (1992): SOS Induction in Escherichia coli by Infection with Mutant Filamentous Phage That Are Defective in Initiation of Complementary-Strand DNA Synthesis. Journal of Bacteriology 174 (5): 1612-1618.
Karlsson F., Malmborg-Hager A.C., Albrekt A.S., Borrebaeck C.A.K. (2005): Genome-wide comparison of phage M13-infected vs. uninfected Escherichia coli. Can. J. Microbiol. 51: 29–35.
Sommer S., Leitao A., Bernardi A., Bailone A., Devoret R. (1991): Introduction of a UV-damaged replicon into a recipient cell is not a sufficient condition to produce an SOS-inducing signal. 254(2):107-17.
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